Living cells typically consume nutrients and oxygen from the surrounding medium, and return metabolic byproducts, including ions, carbon dioxide, lactate, and various proteins, to this extracellular environment. The rate of uptake and excretion of these analytes can provide valuable information regarding the metabolic processes underway inside the cells.
Conventional biological assays inherently exhibit significant limitations. An ideal biological assay is homogeneous (i.e., does not require the introduction of a foreign agent such as a dye), non-invasive (i.e., has no deleterious effect on the biological process), and rapid.
Many tools have been developed to probe the mechanistic processes of cells using internalized reporters such as fluorescent dyes. A device that is able to measure extracellular analytes using a non-invasive, homogeneous assay performed within a container that is compatible with existing invasive tools would be particularly useful.
Some previous approaches relate to oxygen flux rate measurements, since respiration can be deemed to be a basic measure of cell viability. Many devices have been developed to monitor respiration in vitro, through determination of the rate of depletion of oxygen in the extracellular medium. The earliest instruments relied on the change in total gas pressure in a sealed vessel, using the assumption that this change was primarily due to oxygen consumption.
In the 1960s, the Clark electrode (Clark, L. C. Jnr. Ann. NY Acad. Sci. 1962; 102:29-45), and later the miniaturized Clark electrode, enabled a more specific measure of oxygen partial pressure. The relative complexity of the Clark design, and the fact that the electrode itself consumed oxygen, may have hindered its incorporation in a highly parallel instrument suitable for widespread use. However, these devices were deemed successful enough to measure cell viability (Gesinski R M, Morrison J H, Toepfer J R. “Measurement of oxygen consumption of rat bone marrow cells by a polarographic method.” J Appl Physiol. 1968; 24(6):751-754), to profile the toxic effects of drugs and chemicals (Shenoy M A, Biaglow J E, Varnes M E, Hetzel F W. “Inhibition of cultured human tumor cell oxygen utilization by chlorpromazine.” Adv Exp Med Biol. 1983;159:359-68), and to show the effect of agents such as insulin on cellular metabolic processes (Panten U and Klein H. “O2 consumption by isolated pancreatic islets, as measured in a Microincubation system with a Clark-type electrode.” Endocrinology 1982; 111:1595-1600).
More recently, several oxygen sensors have been developed that can enable the design of a non-invasive, homogeneous readout of cellular respiration. Fluorescent compounds, whose response is diminished by the phenomenon of oxygen-quenching, are now available. These compounds can be embedded in an oxygen permeable membrane and exposed to cell media, and can be read using low cost, fiber coupled, semiconductor light sources and sensors (Wolfbeis O S, 2002. “Fiber-Optic Chemical Sensors and Biosensors.” Annal of Chem. 2002; 74:2663-2678).
An ion-sensitive field-effect transistor (ISFET), whose gate region can be exposed to a liquid analyte, has been adapted to measure oxygen pressure using enzyme catalyzed conversion of oxygen (O2) to H+ ions that can be detected by this sensor (Lehmann, M, Baumann W, Brischwein M, Gahle H-J, Freund I, Ehret R, Dreschler S, Palzer H, Kleintges M, Sieben U and Wolf B. “Simultaneous measurement of cellular respiration and acidification with a single CMOS ISFET. 2001.” Biosensors & Bioelectronics. 2001;16:195-203).
Devices have been described and/or demonstrated that incorporate oxygen-quenched fluorophores, ISFETs and other oxygen sensors within sample chambers containing bacteria or mammalian cells for the purpose of measuring respiration rate, viability, or the effect of drugs or toxins. These devices range in size from fluorescent patches attached to the interior wall of large cell culture bottles (Tolosa L, Kostov Y, Harms P, Rao G. “Noninvasive measurement of dissolved oxygen in shake flasks.” Biotechnol Bioeng 2002 Dec. 5;80(5):594-7), to fluorescent sensors embedded within microscopic flow cells fabricated using microfluidics technology (Lähdesmäki I, Scampavia L D, Beeson C, and Ruzicka J. “Detection of Oxygen Consumption of Cultured Adherent Cells by Bead Injection Spectroscopy.” Anal. Chem. 1999; 71: 5248-5252), to microtitre plates with fluorescent compounds suspended within (O'Riordan T C, Buckley D., Ogurtsov V, O'Connor R., Papkovsky D B “A cell viability assay based on monitoring respiration by optical oxygen sensor.” Anal. Biochem. 2000; 278(2):221-227) or deposited upon the wells (Woodnicka M, Guarino R D, Hemperly J J, Timmins M R, Stitt D, Pitner J B. “Novel fluorescent technology platform for high throughput cytotoxicity and proliferation assays.” Journal of Biomolecular Screening. 2000; 5:141-152).
Some patents describe a device for monitoring cells using an oxygen-quenched fluorescent compound that is placed in contact with a broth containing bacteria or mammalian cells. A fluorescence measurement of cells treated with a drug or toxin may be compared to a reference, purportedly to determine the effect of the compound on cellular respiration. In an embodiment, cells are contained within a microplate that is exposed to ambient air. Cells are maintained at a low density in order to maintain viability in this configuration, because high cell density would likely result in anoxia, acidification of the media, and contact inhibition. Measurement times may, therefore, typically be tens of hours or days. In addition, the influx of ambient oxygen and lack of control of sample volume may allow only relative measurement to control to be made. In another embodiment, to limit ambient oxygen influx, mineral oil is placed above the cell media. Because cell density is typically quite low, long measurement times are typically required.
A number of patents and publications describe oxygen flux measurement systems incorporating small, closed sample chambers containing high densities of cells. In these devices, an active perfusion system is used to intermittently restore normal levels of dissolved oxygen, pH, and nutrients. None of these systems are designed or configured to enable the user to easily culture cells, maintain their viability, run experiments in parallel with high throughput, or run other types of assays without detaching and moving the cells.
There have also been approaches to measuring cellular acidification rate. Living cells produce protons (H+ ions) as a byproduct of various metabolic processes, including both aerobic and anaerobic respiration. Protons are also produced when ion exchange pumps on the surface of eukaryotic cells are activated as a result of binding of a ligand with a transmembrane receptor or ion channel. In a fixed volume of extracellular media, this proton flux causes a gradual acidification that can be measured using a pH sensor. Thus, an indication of metabolic rate and/or receptor activation can be determined from a precise measurement of extracellular acidification rate.
A number of pH sensors can be applied to the measurement of cell media. In addition to fluorescent and ISFET sensors similar to those described previously, a light addressable potentiometric sensor has been incorporated in an instrument for rapid measurement of proton flux (Parce W, Owicki J, Kercso K, Sigal G, Wada H, Muir V, Bousse L, Ross K, Sikic B, and McConnell H. 1989. “Detection of Cell-Affecting Agents with a Silicon Biosensor.” Science. 1989; 246(4927):243-247).
One patent describes a device employing a method for measurement of extracellular acidification (pH) as an indicator of cellular metabolism. In this device, a small sample chamber containing a high density of cells is intermittently perfused with media and closed to allow measurement of the pH change resulting from cellular proton excretion. A series of repetitive stop/flow cycles provides kinetic metabolic rate data. Because the sample chamber, once assembled, is fixed in size and contains a high density of cells, active perfusion is required to prevent cell death from the rapid acidification and depletion of oxygen from the media. The addition of a perfusion system to the device results in the need for relatively complex tubing, pumps, and other features, that create cleaning and sterilization problems for the user. In addition, when cells are to be treated with a drug using this device, the drug may need to be perfused over the cells for a relatively long period of time, thereby consuming large quantities of typically scarce and expensive compounds.
Other extracellular analytes can be measured using non-invasive techniques. Carbon dioxide evolution can be determined from the measurement of carbon dioxide (CO2) partial pressure in the media using various fluorescent sensors (Pattison R, Swamy J, Mendenhall B, Hwang C, and Frohlich B. “Measurement and Control of Dissolved Carbon Dioxide in Mammalian Cell Culture Processes Using an in Situ Fiber Optic Chemical Sensor.” 2000. Biotechnology Prog. 16:769-774)(Ge X, Kostov Y, and G Rao. High Stability non-invasive autoclavable naked optical CO2 sensor. 2003. Biosensor and Bioelectronics 18:pp.857-865).
Other ions and chemical constituents can be measured using non-invasive techniques based on optical or semiconductor sensors. In addition, larger molecules such as proteins can be measured using non-invasive techniques that are sensitive to the binding of these molecules to antibodies that are attached to sensors exposed to the extracellular media (Flora K and J Brennan. Comparison of Formats for the Development of Fiber-Optic Biosensors Utilizing Sol-Gel Derived Materials Entrapping Fluorescently-Labeled Proteins. Analyst, 1999, 124, 1455-146).
Other physical phenomenon that support such sensors are surface plasmon resonance (Jordan & Corn, “Surface Plasmon Resonance Imaging Measurements of Electrostatic Biopolymer Adsorption onto Chemically Modified Gold Surfaces,” Anal. Chem., 69:1449-1456 (1997), grating couplers (Morhard et al., “Immobilization of antibodies in micropatterns for cell detection by optical diffraction,” Sensors and Actuators B, 70, p. 232-242, 2000), ellipsometry (Jin et al., “A biosensor concept based on imaging ellipsometry for visualization of biomolecular interactions,” Analytical Biochemistry, 232, p. 69-72, 1995), evanescent wave devices (Huber et al., “Direct optical immunosensing (sensitivity and selectivity),” Sensors and Actuators B, 6, p. 122-126, 1992), reflectometry (Brecht & Gauglitz, “Optical probes and transducers,” Biosensors and Bioelectronics, 10, p. 923-936, 1995) and Wood's anomaly (B. Cunningham, P. Li, B. Lin, J. Pepper, “Colorimetric resonant reflection as a direct biochemical assay technique,” Sensors and Actuators B, Volume 81, p. 316-328, Jan. 5, 2002).
In general, the utility of devices incorporating these sensing technologies for the purpose of measuring secretion of proteins by cells is limited by detection sensitivity. Sensitivity can be increased, typically by increasing cell density in the region proximal to the sensor surface. However, cellular health declines rapidly as cell density increases, due to anoxia, acidification of the media, and contact inhibition. It is possible, but generally undesirable, to adhere cells directly to the sensor surface.
A need exists for the provision of a high cell density for measurement of analytes and a low density for maintenance of cell health and growth. While many devices have been developed for the purpose of measuring flux rates of extracellular analytes, there exists a need to meet requirements that may enable widespread use in the fields of biological research, drug discovery and clinical diagnostics. A need exists for devices with high throughput and ease of use. A parallel configuration may be desirable. Preferably, a tradeoff between long assay times and the length of time to prepare the sample would be eliminated. Lack of these attributes may result in low sample throughput and therefore incompatibility with modern drug discovery and diagnostic activities.
In addition, there is a need for an instrument that can be used to measure extracellular flux rates of cells in a non-invasive manner within a vessel that is commonly used for other high throughput assays, thereby allowing the use of the flux rate measurement as a quality control or complementary measurement to existing assays.
In summary, there is a need for a device that can meet the goals of data quality, compatibility with existing experimental practices, and ease-of-use, thereby enabling widespread adoption of a new technology.